The recent abundance of genome sequence data has necessitated systematic proteomics to decipher the encoded protein networks that dictate cellular function. These include cell-cell interaction, cell activation, cell cycle, signaling pathways, cell proliferation, differentiation, apoptosis, development and many other cellular functions. Initial steps in elucidating the function of an uncharacterized gene product, i.e. a novel protein, often involve studying its interaction with other proteins. Whereas the elucidation of a protein network provides a wealth of information regarding the players, insight into interactions between individual molecules is essential for understanding the contribution of each molecule to a molecular network.
Co-localization studies are often employed to monitor the proximity of one protein of interest to another protein of interest. Co-localization data are also useful as a means of evaluating protein information inferred from genetic data, for instance, supporting or refuting putative protein interactions suggested from for example a yeast two-hybrid analysis. Co-localization of intracellular proteins can be assessed by use of differentially labeled antibodies specifically reactive with endogenous proteins of interest which can be visually detected by fluorescence microscopy via immunofluorescence. Hereto, fixed cells are treated first with a set primary antibodies specific to proteins of interest, and then with a set of secondary antibodies conjugated with fluorescent dye. Antibodies directly conjugated with a fluorescent dye may also be used. Those dyes should differ in excitation and emission wavelength, so that they can be excited independently and observed in separate fluorescent channels. Alternatively, genes encoding the proteins of interest may be fused to a reporter gene encoding a reported protein, like green fluorescent protein (GFP), or tagged with an epitope, such as Myc or HA. Reporters and epitope tags are fused routinely to either the N or C termini of target genes. Dyes specific to membranes or nucleic acids may additionally be used to reveal the cell organelles, for instance, the nucleus can be visualized by DNA staining with DAPI. Images are generally collected on a confocal microscope which ensures that observed proteins are in the same focal plane, and therefore co-localization, if any, is real. Co-localization is revealed by overlap in colors.
However, it should be emphasized that the resolution of confocal microscopy only allows one to detect a global co-localization of proteins and does not necessarily prove close interaction. Overlapping colors do not necessarily imply interacting proteins, i.e. positioning of the proteins within a very short distance, for instance in the range of 3 to 100 Ångstrom. Therefore, co-localization studies routinely require supplementary analysis to investigate whether co-localized molecules represent truly interacting partners. Typical standard biochemical techniques to evaluate putative interacting molecules include co-immunoprecipitation experiments, affinity pull down assays and affinity chromatography.
Co-localization of cell surface molecules, for example, proteins A and B, can also be determined via so-called “patching/capping” experiments. Briefly, upon addition of a multivalent ligand for protein A to viable cells, a clump or patch of protein A molecules assembles in the membrane. If the cell is alive and metabolizing actively, patches are formed which can further assemble into a cap in a process called “capping”, preferably occurring at 37 degrees Celsius. The patches/cap can be stained by indirect fluorescence staining procedures. Subsequently, cells may be counterstained at 4 degrees Celsius (to retain the patches/cap) with a dye-conjugated antibody against protein B to evaluate whether protein B has moved together i.e. is co-localized with the “patched/capped” protein A or whether protein B is still diffusely distributed on the cell surface.1 Although this method works in practice for assessing co-localization of surface membrane proteins, it is time consuming, needs experience and can only be evaluated by dye microscopy, not by flow cytometry. Furthermore, microscopic procedures are not suitable as a high throughput method for the evaluation of interacting molecules.
A particularly elegant method to detect closely interacting molecules involves fluorescence resonance energy transfer (FRET). In FRET, a dye (called a “donor”) transfers, after excitation by a light source, its energy to another dye (called “acceptor”). The energy transfer occurs when the emission spectrum of the donor dye overlaps significantly with the excitation spectrum of the acceptor. Sufficiently close juxtaposition of the two dyes, generally closer than 100 Ångstrom, but preferably closer than 50 Ångstrom, is essential for energy transfer between the donor/acceptor pair. One Ångstrom, a metric unit of length, is equal to 0.1 nanometer or 10−10 meter. FRET is usually based on the interaction between donor and acceptor dyes that are both fluorescent. However, FRET can also be detected by the quenching of donor fluorescence using a nonfluorescent acceptor dye. Nonfluorescent acceptor dyes are in general advantageous because they eliminate the background fluorescence that results from direct acceptor excitation. In the present invention, it is possible to monitor juxtaposed probes on interacting molecules using a fluorescent donor dye and a nonfluorescent acceptor dye. Specific binding of a set of probes to non-interacting molecules will give a basal fluorescence signal. Upon close interaction of the molecules, FRET between the probes will quench the donor fluorescence. Rather than measuring an increase in acceptor fluorescence, use of a nonfluorescent acceptor involves measuring a decrease in donor fluorescence. Generally, detection of a decreased signal is less sensitive compared to detection of an increased signal. Therefore, a method according to the invention is preferably practiced using a fluorescent donor and a fluorescent acceptor dye.
FRET energy transfer efficiency is inversely proportional to the sixth power of the distance between the donor and the acceptor. FRET, first described by Förster, has become extremely important for modern cell biology because FRET allows to measure distances between molecules on a scale of a few nanometers. This is far below the resolution limit of modern optical far field microscopy, which currently is at approximately 100 nm. FRET technology has been used for detection of various individual (bio)molecules. For example, U.S. Pat. No. 6,235,535 discloses a fluorescence-based immunoassay method for the detection of an analyte in a biological sample. The method is based on the ability of a multivalent analyte (antigen) to induce aggregation of identical receptor molecules (antibodies) labeled with a fluorophore, which molecules are immobilized onto yet freely mobile on a lipid membrane. Antigen-induced aggregation of the receptors causes FRET to take place. Also in U.S. patent Publication 2002/0081617, antibodies directed to the same epitope but labeled with either a donor of acceptor dye are immobilized, in this case onto beads. Upon addition of an analyte (antigen) of interest, the analyte functions as a bridge and brings a pair of antibodies into close proximity of each other which leads to FRET. Thus, U.S. Pat. No. 6,235,535 and U.S. patent Publication 2002/0081617 both relate to the detection or measurement of an analyte using immobilized, dye-conjugated probes and FRET-based detection methods. Since the probe sets of U.S. Pat. No. 6,235,535 and U.S. patent Publication 2002/0081617 are directed to a single molecule or molecular epitope, they are essentially not suitable for detecting distinct interacting molecules.
The extreme sensitivity of the FRET process on the distance between molecules renders it a very useful tool for the resolution of intracellular protein arrangements and protein dynamics. The presence of FRET indicates intermolecular interaction since it is observable only for nanometer-scale fluorophore distance. This implies in particular that simple co-localization of two molecules, e.g. proteins, is not sufficient to yield energy transfer. FRET is a technique that can give clear, unambiguous answers to questions about protein-protein interactions. FRET measurements can be used to determine protein interactions at the cell surface2. The “green revolution” initiated by the introduction of the green fluorescent protein (GFP) from Aequorea victoria and the later developments of GFP-mutants possessing different spectral properties offered the possibility of simultaneous expression of different proteins, artificially tagged with fluorescent donor and acceptor domains in the same cell.3,4 This allowed measurement of their interactions by FRET. The combination of Cyano Fluorescent Protein (CFP) (donor) and Yellow Fluorescent Protein (YFP) (acceptor)—tagged proteins is often used. This FRET pair can be used to monitor the proximity of the two attached fluorescent tags in 3-6 nm. Co-expression of CFP- and YFP-tagged proteins has been successfully used to analyze short time changes in protein-protein interactions, e.g. oligomerization, co-localization, complex formation, activation of kinases and mapping of enzyme activities in living cells. FRET technology was also used in a highly specific fluorescence lifetime imaging microscopy (FLIM) method for monitoring epidermal growth factor receptor (EGFR) phosphorylation in cells. EGFR phosphorylation was monitored using a GFP-tagged EGFR and Cy3-conjugated anti-phosphotyrosine antibodies.5 
Although fluorescently tagged proteins have proven to be very useful, they do have limitations, such as their significant size (>200 amino acids). Also, the overall folding and tertiary structure of a tagged protein may be different from that of the native, non-tagged protein. This may result in different, erroneous interactions with other molecules. Drawing conclusions with respect to the addressed protein-protein interactions on the basis of FRET data between pairs of tagged proteins, as well as performing comparisons with normal cellular functions in living cells, is justified only if the recombinant, tagged proteins behave similar to the corresponding endogenous wild-type proteins. For instance, the expressed fluorescently-tagged protein should reveal the same intracellular distribution as the wild-type protein; the expression of the tagged protein per se should not induce or inhibit cellular functions and the tagged protein expressed in the cell should not create significant background FRET-signal, for example, due to overexpression, pH shift etc. Another major drawback of the use of recombinant, tagged proteins lies in the fact that it requires transfection or co-transfection of a chimeric construct or constructs of interest into a cell and selection of a cell showing adequate expression of a construct to yield a functional protein. Such a system does not allow detection of an endogenous protein and can therefore not be used to evaluate endogenous interacting molecules. One report describes the use of FRET technology to monitor ligand-induced dimerization of an endogenous cell surface receptor by means of a receptor-specific antibody that was directly conjugated to either the donor dye FITC or the acceptor dye Cy3.6 The ability of a ligand to induce receptor dimerization was assessed by flow cytometric analysis of FRET between FITC and Cy3. A pair of antibody conjugates was used to study cell surface proteins on human lymphocytes: the CD8alpha chain detected by pairs of antibodies against different epitopes; the very late antigen 4 (VLA4), a heterodimeric alpha4beta1 integrin, was detected via FRET between antibody conjugate pairs specific for either integrin beta1 (CD29) or integrin alpha4 (CD49d); association of T-cell receptor (TCR) with a soluble antigen ligand was detected by FRET when anti-TCR antibody and MHC class I/peptide complexes were used. In yet another report, antibody mediated FRET technology was used to measure the interaction of c-kit receptor with its ligand SCF (stem cell factor).7 Thus far, antibody-mediated FRET technology has not been applied to detect intracellular protein-protein interactions.
FRET technology has also been applied for the detection of a protein-DNA interaction on the basis of a so-called indirect binding principle. For example, it was used to monitor the interaction between the p65 subunit of the transcription factor, NF-kappaB and its DNA binding site. NF-kappaB is of great relevance to the pharmaceutical sector due to its ability to regulate a number of genes involved in various immune and inflammatory responses. As such NF-kappaB has been implicated in several disease states including various viral infections (HIV), arthritis and cancer. An anti-GST antibody labeled with Cy3 (approx. 7-12 dyes per molecule) is allowed to interact with an affinity purified GST fusion protein of p65 and GST (p65GST). A double-stranded DNA (dsDNA) sequence which contains the NF-kappaB binding site was singly labeled with Cy5 at the 5′ end of the coding sequence. This was then incubated with a Cy3 labeled antibody and p65GST. The reaction was done either in the presence or absence of unlabeled non-specific or specific competitor dsDNA. In the absence of either competitor, binding by p65-GST resulted in FRET between the Cy3 donor molecules on anti-GST and Cy5 acceptor molecule on dsDNA.
Thus, it would be advantageous to possess a method that allows the detection of interactions between endogenous intracellular proteins and/or other molecules such as nucleic acids, lipids and/or carbohydrate moieties. Particularly challenging is the detection of intermolecular interactions at the single cell level.